Stereotaxic Surgery
The stereotaxic frame is designed to hold the animal’s head perfectly still, and in a reproducible way such that a brain target can be reliable located in 3-D space with reference to some measurable location. The frame allows for manipulation of the probe in an anterior-posterior direction, lateral direction and dorsal-ventral direction. An implant can be placed on an angle from vertical, and the head can be tilted up or down from. Co-ordinates should be reported with reference to some structure, nose bar level, and angle of implant. All will affect the accuracy of the surgery. Typically, bregma is used as an initial landmark on the skull. The skull of a rodent is formed by four bone plates, each separated by sutures (interface of the edges of two bone plates.) There are two parietal bone plates, separated by the sagittal suture, a frontal plate and an occipital plate. Bregma is the point at which the frontal plate meets the anterior edges of the parietal plates along the sagittal suture (i.e., where the coronal suture meets the sagittal suture). The intersection of the sagittal suture with the suture along the anterior edge of the occipital plate (lamdoidal suture) is known as lambda. Areas of the brain are typically defined as being x mm anterior/posterior to bregma, and y mm lateral to midline. For depth, the target is either z mm below the skull surface or dura. Dura is the more accurate of the two, as skull thickness is more variable than the size of the brain. Care must be taken not to damage dura when drilling through the skull. If dura is destroyed, skull surface can be used, and the target depth can be modified by 1 to 1.5 mm for rats, and 0.6 to 1 mm for hamsters. These numbers will depend on the age of the animal and the site of the burr hole. These adjustments should be modified based on experience. A surgeon should be able to collect data from surgeries in which dura was not damaged, and thus determine the mean thickness of the skull in a given cohort of animals at a certain site. Alternative reference points are lambda, interaural zero and a removable zero bar. In the latter two cases, the head of the animal must be perfectly centered relative to the frame. If you are going to use interaural zero as a reference, this must be measure before the surgery. Place the ear bars close together and at the same lateral position and then p-lace your probe precisely between the tips of the ear bars. The position of the nose/tooth bar should be specified (x mm above or below interaural level). The fulcrum of the head in the stereotaxic frame is around the interaural line, and the angle of the head is determined by the position of the nose bar. Typically the head is leveled, that is, it is positioned such that bregma and lambda are at the same dorsal/ventral position with reference to the frame. This position is typically 2 mm below the interaural level for hamsters and 3.3 mm below the interaural level for rats. Always report whether you leveled the head, or if not, at which level the nose bar was set. Stereotaxic co-ordinates are meaningless without information on the tilt of the head. When doing stereotaxic surgery, it is often handy to have a riser of some sort for smaller animals. We use a 1 cm tall block wrapped in paper towel for hamsters. Towel should be place under a rat so that its body is not in contact with the metal base of the frame. A heating pad that can fit inside the stereotaxic frame would be even better than paper towels, as it is even better as it actively keeps the animal warm.
Vernier Scale Precise measurements are accomplished with the stereotaxic frame by the use of Vernier scales that allow for readings as precise as 0.1 mm to be made. One side of the scale (located on the moving bar in each case) is marked off in centimeters (numbered) and millimeters (unnumbered, but marked). Most co-ordinates for rodents are reported in millimeters, and thus you should record your measurements this way (e.g., 4.25 cm = 42.5 mm). There is a second scale opposite the first and attached to the non-moving aspect of each scale. This second scale is divided into 10 equal units, and the whole scale spans 9 mm. To read the Vernier scale, locate the zero on the small scale. Determine which mm division on the large scale it is in front of. For example, if it is between the third and fourth small division after the 4 cm mark, this would be 4.3 cm or, as you will record, 43 mm. To determine the fraction of a millimeter for the given measurement, the number will come from the small scale. It will be the number of the line on the small scale that lines up perfectly with a line on the large scale. This is very important. The tens and ones measurements are determined by the position of the small scale’s zero, and are read from the large scale. The tenths position is determined by finding a line on the small scale which lines up perfectly with a line on the large scale, and is read from the small scale. There currently is a web site with a java applet which simulates a Vernier scale. I highly recommend practicing reading the Vernier scale, as misreading it will lead to a misplace surgery. The URL is: http://www.upscale.utoronto.ca/PVB/Harrison/Vernier/Vernier.html
Ear Bars The first step in stereotaxic surgery, once the animal has been prepared, is to place it in the ear bars. This is probably the hardest step in stereotaxic surgery, and also one of the most important steps. If the animal is not place in the ear bars correctly, the co-ordinates will not be accurate. Proper and repeatable placement is essential for accurate stereotaxic surgery. There are two main types of ear bars for rat or hamster surgery; the standard rat ear bars Kopf model # 957), and the atraumatic ear bars (either Kopf model # 951, or # 955). The standard ear bars are quite pointy, and are the easiest to place in the ears reliably. Unfortunately, they frequently damage the tympanic membrane and the bones of the middle and inner ear. This can lead to some balance disturbances or middle ear infections. The ear bar should be disinfected between each use in case damage is done, so that infectious agents are not introduced to the middle ear. The atraumatic ear bars come in a variety of forms, either similar to the standard, yet more blunt, or tipped with a wide point. These ear bars are less likely to cause damage, yet they are error prone. It is substantially more difficult to place the animal accurately and reliably in these ear bars, and the head is prone to slipping out of them. Essentially, you should gauge the needs of the surgery at hand. If the need is simply to hold the head still so that a platform can be fixed to the skull, the atraumatic ear bars are best. If precision is essential, the standard ear bars can be used with extreme care to avoid damaging the tympanic membrane. Placing the animal in the ear bars is difficult. It is difficult to see what is being done. Essentially, the left ear bar should be fixed in place at its anticipated final position. Holding the animal in your right hand, place it such that the tip of the bar is inside the ear canal opening. Reach around with your free hand and grasp the head, thumb on top, index and middle fingers on the lower jaw. Lay the body down so as to free your right hand of its bulk and work only with the head. Holding the head level, it should be drawn up and back slightly, allowing the tip to fit into the ear canal which goes into the skull of the rodent on a downward and forward angle. When this bar is in place, rotate the nose ever so slightly towards the left side of the frame, this will place the free ear canal at an angle such that the second ear bar can clear the external auditory meatus and slide easily into place in the ear canal. As then ear bar in inserted past the outer ear, rotate the nose back to midline. This procedure for the second ear is most easily accomplished if the second bar is just loose enough to slide horizontally without much vertical movement. When placed correctly, the head should be able to swing up and down quite easily, and there should be no lateral movement of the nose. However, a little lateral movement may merely indicate that the ear bars are not in far enough, rather than indicating that you have missed. Each of the ear bars should be close to the same lateral position (as measured by their individual scales), although this is only necessary if an external zero point is being used, such as a removable zero point.
Nose / Incisor Bar The nose bar help stabilize the skull. It is important that the skull be held firmly in place for two reasons. First, a loose head moves, and it is impossible to measure coordinates accurately on a moving head. Second, a moving head will vibrate more in the ear bars and nose bar. If this occurs, there is the risk of unnecessary damage to the animal, in the form of ear trauma or damage to the upper palate and teeth. Do not worry about cutting air flow by clamping down too hard with the bar over the nose, as the nasal cavity is within the skull, not between the skull and the skin. However, do not tighten it so hard that there will be tissue and bone trauma. You should not be able to slide a thin spatula between the bar and the animal’s snout. To place an animal in the tooth / nose bar (Kopf model #920), first correctly place it in the ear bars. Using a spatula, open the mouth but gently pushing down on the lower jaw. Slide the tooth bar forward until it is just under the top incisors. Rest the top incisors on the tooth bar, and press the lower jaw and tongue down. With this done, it should be possible to slide the tooth bar between the upper and lower incisors. Carefully push the tooth bar into the mouth and let the top incisors fall through the hole in the tooth bar, then gently push the tooth bar away from the animal until the inside edge of the tooth bar hole touches the inside surface of the top incisors. Without letting go of the tooth bar at this point, tighten the tooth bar so that it no longer moves back and forth. If you let go first, it will move forward from the weight of the animals head, and will not end up in the correct location, potentially altering the tilt of the head, and thus invalidating your surgery coordinates. A note of caution, be careful moving the tooth bar into or out of the mouth. The incisors must be clear, or you will break them. Always make slow controlled movement and watch what you are doing. One the animals head is fixed in place, establish your sterile field and prep both yourself and the incision area for surgery. This includes washing your hands, donning sterile surgical gear (hair and face mask, gown, gloves), draping the animal in sterile drapes and disinfecting the surgical site. Draping the animal is challenging when it is in a stereotaxic frame, so consult your veterinarian for advice on an acceptable and appropriate technique.
Implantation
Using the stereotaxic frame, it is possible to place an implant in a precise location in the brain. An excellent protocol with video is available (Geiger et al., 2008; http://www.jove.com/index/Details.stp?ID=880). An excellent guide is also available (Cooley and Vanderwolf, 1990). A variety of implants are used in behavioral neuroscience. The two most common would be an electrode and a cannula. A cannula is a tube which allows material to be removed from, or delivered to, a specific area of the body. For neuroscience research, an indwelling cannula can be positioned such that a very small amount of drug can be delivered to a discrete region of the brain. Commercially available cannulas and electrodes are available from Plastics One (Roanoke, VA). There are three components to a cannula. First, there is the implanted guide cannula. Typically, this should be implanted such that its tip is some distance from the target structure (we typically use 1 mm). A dummy cannula or dummy cannula is placed in the guide cannula after surgery and between injections. This prevents fluid from seeping into and thereby clogging the guide cannula. The third component is the injection cannula, which has a collar to prevent it from proceeding too far into the brain. Both the dummy and injection cannula obtrude from the end of the guide cannula by 1 mm. This is essential, as if they were to end flush with the guide, it is likely that the fluid would flow back up the cannula. If they were to obtrude by more than 1 mm, a small bend or kink in the injection cannula could lead to it missing its target. The dummy cannula should be the same length as the injection cannula, so that tissue near the target structure is not damaged for the first time when doing your experimental injection. Before placing the cannula in the holder on the stereotaxic frame, place a small stylet (wire with a hook) into the cannula to fill the space inside the cannula during surgery. Cannulas may need to be cut to length before the surgery, and can have their tips beveled to aid passage through the brain tissue. Electrodes may need preparation prior to implantation. Insulation may need to be removed, and the tips may need to be separated. They may also need to be cut to length, in which case the insulation should be examined to ensure that it has not cracked. Once the animal is placed in the frame, make an incision (see above). The fascia (connective tissue) can be blunt-dissected with cotton swabs. Place two swabs close together, press down firmly and pull them apart, laterally a few times, and rostral / caudal a few times. The skin can be retracted with hemostats (rats) or bulldog seraphines (hamsters) by clamping them onto the fascia. Avoid clamping the muscle or the skin, as this will produce extraneous tissue damage and impair healing of the wound. For indwelling electrodes and cannulas, room must be created on the skull for the head cap that will be created. This is very important when the subject is a hamster, as they have quite small skulls compared to a rat. This can be done with a metal spatula, or the soft shaft of a sterile cotton swab. Start along midline at the anterior end of the incision and move laterally until you catch the edge of the muscle. With firm controlled pressure, ease the muscle off of the skull. Work your way toward the posterior end of the incision. Only move what you need to, as this muscle is what the animal uses to raise its lower jaw. It is attached vertically at the caudal portion of the skull, so the animal will still be able to eat, even with this removed, but there is no sense in making it any more difficult for the animal than necessary. This may not be necessary for rats, depending on how medially the implant will be placed. Once the incision is retracted, and the muscle prepared, dry the skull with a cotton swab that has been dipped in 70% alcohol. This will dry the skull quite quickly, making the suture extremely easy to locate. Use a metal spatula to quickly and firmly scrap any bleeding points on the skull. This will clog the small openings with bone, and stop the flow of blood. It is essential that the skull be clean and dry. Mount your implant in the holder on the stereotaxic arm. Manipulate the arms to place the tip of your implant on bregma (if that is your landmark). This is often easier to do with the aid of a small jeweler’s magnifying glass or stereomicroscope. Record its location, and calculate your target. Raise the implant up sufficiently far so that it will clear the skull (1 or 2 mm), and move the cannula to its A/P and lateral targets (a burr hole must be drilled before the D/V coordinate can be reached). Record the D/V coordinate of the skull, even if dura is the landmark for the D/V coordinate. Raise the implant up a bit and make a mark on the skull with a pencil or a fine tipped waterproof marker. Raise the implant up high enough so that it will clear the edge of the wound and move it laterally to get the horizontal bar of the frame out of your way. The head cap will be secured to the skull by placing 4 jeweler’s screws into the skull. The heads of these screws will be imbedded into the dental acrylic. Mark 4 point for the screws, and ensure that they are sufficiently far apart from one another and the implant target such that each screw head will not touch another or the implant. If possible, have screws to either side of the implant, both in front and behind. Do not place a screw on a suture or near midline. There is a blood vessel known as the sagittal sinus directly below the sagittal suture. If this sinus is damaged, there will be a profuse amount of bleeding. Using a drill bit slightly smaller than the screws, drill a tiny hole for each screw. Angling the burr holes towards the center of the brain will give the final head cap more stability than if the holes are precisely vertical and parallel. Before your first attempt to drill, practice touching the skull with the drill while it is not turned on. Become aware of the sensation of touching the skull. Try raising it and lowering it a few times, making very small controlled movements. Repeat this with your eyes closed, and really concentrate on the feedback. This is the sensation that you will try to maintain during drilling. Practice small movement, as the skull is only about 1 mm thick. When drilling, press down only hard enough to maintain that sensation, and when you feel it give a little, withdraw the drill. To drill, hold the drill in your dominant hand and support your wrist of hand with your other hand. Each hole should be drilled so that it is tilted slightly towards the center in such a way that all the screws point in separate directions. This will make it more difficult for them to come out. For hamsters and rats, there will be two stiff points in the drilling, one at the outer surface of the skull, and one at the inner surface of the skull. There is a soft section of bone between these two layers. You may feel the drill give even when you have not completed the hole. This sensation would be from entering this soft layer of the skull. Before placing the screw, ensure that you are through both layers. This can be done by probing the burr hole with the drill when it is not turned on. If minimizing damage to the cortex is critical to your surgery, shallow holes and screws can be used that do not completely pass through the skull. The head cap will be weaker and may come out, but the cortex will be intact. As you have a choice of where the screws will go, but don’t have a choice as to the location of the burr hole for the implant, it is recommended that the screw holes be drilled first, in case the cannula hole must be near a location which is prone to bleeding. Should you get a lot of bleeding when drilling the burr hole, you will at least have already completed the task of implanting the screws. Once all four screw-holes are drilled, the screws can be placed in the holes using a small pair of curved forceps and a small jeweler’s screwdriver. The head should not be flush with the skull, as it needs to be embedded in the dental acrylic. However, the majority of the shaft should be in the skull, as the further it is into the skull, the better it will hold. To drill the implant burr hole, lower the drill until it touches the skull. Then, without much further downward pressure, work in a circular motion, carving a hole sufficiently large to accommodate the implant without having it touch the sides at all. This fine drilling is often accomplished more easily with the aid of a small jeweler’s magnifying glass or stereomicroscope. Carve away the bone until you can see the dura. When the burr hole is drilled, return the implant to it lateral and A/P coordinates, and assess whether or not it will fit into the hole without touching the sides. It is essential that it not touch the sides, as this will cause a small deflection, which translates into an increasingly large error the deeper the implant is inserted. Once it has been determined that the implant fits freely into the burr hole, lower it down into the hole until it makes contact with dura. A small depression in dura will be observed. Stop here, and record the DV coordinate if dura is being used as a landmark for depth. Prior to inserting the implant to its D/V coordinate, raise the implant up a bit and puncture dura with a sterile sharp needle. This is necessary as dura is quite tough. If it were not punctured, it would stretch and break and cause a lot of trauma in the process. If a fine electrode were being implanted, it is more likely that the electrode would bend rather than dura break, which may ruin the electrode, and would most certainly ruin your coordinates that were based on a straight electrode. Once dura has been punctured, stop any bleeding with cotton swabs. Dry the skull and clean it of blood and bone chips. This is essential to ensure that the dental acrylic forms a good firm bond with the skull. Proper clearing of bone chips is also essential for the animal’s comfort, as if these remain in the wound, they will irritate the animal. After this, lower the implant to its target slowly (a speed of 1mm per 10 seconds is good). Do not lower it too fast or you may overshoot your target. Once the implant is in place, clean the skull again, being careful not to touch the implant. A Kim-wipe, twisted to a point between your gloved hands, can be used to soak up any fluid under the screws and implant. The dental acrylic (Dentsply repair caulk) should be applied in three coats. The first coat should be quite runny, and must be applied quickly. Mix the acrylic with its solvent in a Petri dish. Do not mix the acrylic much (tilting the dish a couple of times is sufficient) as this will start the curing process, making the acrylic more difficult to work with. Using the pointy end of a metal spatula, apply the acrylic generously to the area between the screws and cannula. This runny coat will fill all the crevices on the skull, and form a solid hold. Shape it with the long edge of the spatula, scooping it towards the center of the skull and away from the tissue. Allow it to cure a little bit to make this task easier. Allow it to dry such that it will not dent when poked with the spatula. Once it is dry, apply a larger, less runny coat. This coat will form the body of the head cap and should come up the sides of the cannula about half way. Ensure that the holder is not embedded in the acrylic, or you will be unable to remove it at the end of the surgery. Also ensure that enough of the implant is above the acrylic such that the dummy cannula or dust cap may still be threaded all the way onto the guide cannula. When this coat is dry, remove the implant-holder. This will give you better access to the head cap for the final coat. This final coat should be extremely runny (mostly solvent). It should be dabbed on in small amounts where the head cap is rough. It should not be shaped, as this will create more rough edges. The purpose of this coat is to smooth out the head cap so that it will not irritate the animal. A rough spot will cause such irritation and the animal will groom at the spot, possibly opening the wound and introducing infection. While the final coat is drying, the seraphine clips or hemostats holding the wound open can be removed. The tissue can be rehydrated with generous amounts of saline. A 5% lidocaine solution can be applied to the skin to minimize pain during suturing. When the head cap is dry, the wound can be suture (as described above) with sutures both rostral and caudal to the head cap. Ensure that the wound is loose enough around the head cap that should it come detached from the skull, that it will be able to fall out of the head, rather than be held in the head in by the skin. If it can not come out when detached, the implant would move around inside the brain, causing massive damage to the central nervous system. Prior to removing the animal from the ear bars, remove the stylet inserted during surgery, and replace it with the proper dummy cannula for cannula implants, and simply attach a dust cap for electrode implants. Loosen the nose bar, and carefully remove the tooth bar. Then, while holding the scruff off the animal, loosen and remove one ear bar. This is sufficient to allow you to remove the animal. Wrap it in paper towel to aid in thermoregulation and place it in a warm cage near a heat lamp. When it has recovered enough to be mobile, give it an injection of an opiate, such as buprenorphine. Follow-up with proper post-operative care and monitoring.
Devascularization Lesion of the Forelimb Area
The Devascularization Lesion, or pial strip, is a simple model of stroke. The procedure for producing a stroke of the forelimb portion of the motor cortex is as follows:Anaesthetize the rat.
Shave the top of the rat’s head.
Place the rat in the stereotaxic frame.
Disinfect the scalp.
Inject lidocaine along the incision line
Incise the scalp along midline, and retract the tissue.
Mark an area of the skull on the side contralateral to the preferred forepaw. This area will run from 4 mm anterior to bregma to 1 mm posterior to bregma and from 1–4 mm lateral to midline.
Using a dental drill, shave down the skull in the area marked. Since dura is adhered to the inside of the skull, simply cutting out a bone flap would rip dura and lead to a lot of bleeding. Carefully shaving away the skull minimizes the damage, and makes it easier to see what you’re doing.
Use a fine needle (26-30g) and bend the tip to about 60 degrees. Use this to puncture dura in an area with little vascularization. Next, use some fine forceps to lift dura at the puncture. Then, using fine spring scissors, carefully remove the dura exposed by the bone-window.
With a sterile saline-soaked cotton swab, gently rub away the vasculature over the cortex.
Fill the bone window with sterile, saline-soaked, gelfoam to protect the brain from further damage.
Close the incision with sutures or wound clips.
Place the rat in a proper recovery location and provide appropriate analgesics and post-operative care.
Stereotaxic Injection – 6OHDA
Neurotoxic lesions are useful models for understanding how specific brain regions regulate behavior. Neurotoxic lesions allow you to eliminate specific cell-types, or to lesion cell bodies but leave fibers of passage intact. The following section will describe the procedure for producing a lesion of dopaminergic cell fibers in the striatum as a model of Parkinson’s Disease. This lesion protocol is based on that described by Ben et al. (1999). Prepare the neurotoxin 6-hydroxydopamine (6-OHDA) fresh just before the surgery. Prepare it to a concentration of 4 μg (salt weight) / μl in sterile physiological saline containing 1% ascorbic acid. Four 2 μl infusions will be performed, so you will need at least 8 μl. Control rats should receive infusions of 2 μl of physiological saline containing 1% ascorbic acid at the same coordinates (sham lesion). Unilateral lesions can be used if the deficits produced by the bilateral lesions are too severe to permit behavioral testing. See the “Implantation” section above for more details on stereotaxic surgery.
Anaesthetize the rat.
Shave the top of the rat’s head.
Place the rat in the stereotaxic frame.
Disinfect the scalp.
Incise the scalp along midline, and retract the tissue.
Clean the skull so that bregma can be visualized.
Mount a Hamilton microsyringe in the stereotaxic frame using a microsyringe holder.
Manipulate the stereotaxic arms to place the tip of the microsyringe over bregma, and not the coordinates for bregma.
Injections will be placed at the following locations relative to bregma and skull surface:
Site 1
Site 2
Site 3
Site 4
AP
+1.7 mm
-0.92 mm
+1.7 mm
-0.92 mm
ML
+2.8 mm
+4.0 mm
-2.8 mm
-4.0 mm
DV
-5.6 mm
-5.5 mm
-5.6 mm
-5.5 mm
Mark the four locations on the skull and using a dental drill make 4 small burr holes large enough to accommodate your microsyringe.
Using a small sterile syringe tip, puncture dura to facilitate entry of the microsyringe.
Slowly lower your microsyringe into position, descending ventrally at about 5mm/min.
Infuse at a rate of 0.5 μl/min. When each infusion is in complete, leave the microsyringe in place for an additional minute to aid infusate diffusion.
Fill the burr holes with sterile bone wax.
Hydrate the incision tissue with sterile saline containing 5% lidocaine.
Close the incision with sutures or wound clips.
Place the rat in a proper recovery location and provide appropriate analgesics and post-operative care. Depending on the severity of the lesion, the animal may have pronounced motor impairments, so careful monitoring of eating and drinking is critical for the days following the lesion.
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